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Biochemistry. Author manuscript; available in PMC Nov 23, 2011.
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PMCID: PMC2982868
NIHMSID: NIHMS242824

Lipoic Acid Synthesis: A New Family of Octanoyltransferases Generally Annotated as Lipoate Protein Ligases

Abstract

Bacillus subtilis lacks a recognizable homologue of the LipB octanoyltransferase, an enzyme essential for lipoic acid synthesis in Escherichia coli. LipB transfers the octanoyl moiety from octanoyl-acyl carrier protein to the lipoyl domains of the 2-oxoacid dehydrogenases via a thioester-linked octanoyl-LipB intermediate. The octanoylated dehydrogenase is then converted to the enzymatically active lipoylated species by insertion of two sulfur atoms into the octanoyl moiety by the S-adenosyl-L-methionine radical enzyme, LipA (lipoate synthase). Bacillus subtilis synthesizes lipoic acid and contains a LipA homologue that is fully functional in E. coli. Therefore, the lack of a LipB homologue presented the puzzle of how B. subtilis synthesizes the LipA substrate. We report that B. subtilis encodes an octanoyltransferase that has virtually no sequence resemblance to E. coli LipB, but instead has a sequence that resembles that of the E. coli lipoate ligase, LplA. Based on this resemblance these genes have generally been annotated as encoding a lipoate ligase, an enzyme that in E. coli scavenges lipoic acid from the environment, but which plays no role in de novo synthesis. We have named the B. subtilis octanoyltransferase LipM and find that, like LipB, the LipM reaction proceeds through a thioester-linked acyl enzyme intermediate. The LipM active site nucleophile was identified as C150 by the finding that this thiol becomes modified when LipM is expressed in E. coli. The level of the octanoyl-LipM intermediate can be significantly decreased by blocking fatty acid synthesis during LipM expression and C150 was confirmed as an essential active site residue by site-directed mutagenesis. LipM homologues seem the sole type of octanoyltransferase present in the Firmicutes and are also present in the Cyanobacteria. LipM type octanoyltransferases represent a new clade of the PF03099 protein family suggesting that octanoyltransfer activity has evolved at least twice within this superfamily.

Keywords: Lipoic acid synthesis, Bacillus subtilis, octanoyltransferase

INTRODUCTION

Lipoic acid is a covalently-bound enzyme cofactor required for energy-conserved decarboxylation of 2-oxoacids in aerobic metabolism as well as for glycine cleavage in all three domains of life. In each lipoic acid-dependent enzyme a specific subunit is modified by attachment of lipoic acid to a specific lysine residue located within a highly conserved domain or domains (called lipoyl domains or LD). An amide linkage is formed between the carboxyl group of lipoic acid and the ε–amino group of the specific lysine residue. During catalysis the protein-bound lipoamide moieties serve as carriers of reaction intermediates among the multiple active sites of these multienzyme complexes.

There are four known 2-oxoacid dehydrogenases. These are pyruvate dehydrogenase, 2-oxogluatarate dehydrogenase, branched chain dehydrogenase and acetoin dehydrogenase (1), although many organisms have only a subset of these enzymes. 2-Oxoglutarate dehydrogenase is a component of the citric acid cycle whereas pyruvate dehydrogenase is required for entry of carbon into the cycle and thus both enzymes are required for aerobic respiration. The branched chain dehydrogenase is required for branched chain fatty acid production in bacteria and also for degradation of branched chain amino acids. Acetoin dehydrogenase degrades the fermentative product, acetoin (3-hydroxybutanone) to acetyl-CoA (2).

Most of our knowledge of lipoic acid biosynthesis comes from studies in E. coli (3) (Figure 1). In the first step of the pathway an octanoyl moiety is transferred from the fatty acid synthetic intermediate, octanoyl-S-acyl carrier protein (ACP) to a specific lysine moiety of the LD by an enzyme called LipB or octanoyl transferase (systematic name; octanoyl-[acyl-carrier-protein]-protein N-octanoyltransferase; EC 2.3.1.181). The LipB reaction proceeds via an octanoyl-thioester enzyme intermediate (4). Once the lipoyl-dependent complex has been octanoylated, the LipA lipoyl synthase (EC 2.8.1.8) inserts two sulfur atoms, the first at octanoate carbon 6 and then at carbon 8 to yield the dihydrolipoyl-LD which is oxidized to lipoyl-LD (5). LipA is an S-adenosyl methionine radical enzyme and the source of the sulfur atoms is a LipA iron-sulfur cluster (6). Thus, in this unusual pathway lipoic acid is synthesized on its cognate enzymes. E. coli also has an enzyme called LplA that functions to scavenge lipoic acid from the environment (7, 8). LplA is a lipoate protein ligase (EC 2.7.7.63) that first activates lipoate with ATP to give enzyme-bound lipoyl-adenylate (9). This intermediate is then attacked by the ε–amino group of the LD lysine residue to give the lipoylated protein. E. coli. LplA is also active with octanoic acid (10) and can bypass the LipB step of the biosynthetic pathway in the presence of exogenous or endogenous octanoic acid (11, 12). Despite the lack of sequence similarity and the fact that they catalyze reactions that proceed by markedly different chemistries, LipB and LplA have similar three-dimensional structures (13) as first predicted by Reche (14). E. coli LplA and M. tuberculosis LipB can be aligned over the length of LipB (the smaller protein) with a Cα value of 2.5Å. Together with the biotin protein ligases, these enzymes form the highly divergent PFAM 03099 protein family. Despite the sequence divergence, the cofactor ligands of the LipB, LplA and the E. coli BirA biotin protein ligase crystal structures are in register indicating that the substrate binding geometries are similar (13).

Figure 1
Octanoylation of Lipoyl Domains

Although Bacilli have an unusually large number of lipoic acid-dependent enzymes (all of the enzymes listed above), the annotations of these genomes argue that these bacteria have an incomplete lipoic acid synthesis pathway. Although Bacillus subtilis encodes a readily recognized and functional LipA homologue (15), as well as multiple putative LplA homologues, no LipB homologue can be recognized. Hence, it was unclear how this organism produces the octanoyl-LD substrates required for sulfur insertion by LipA. A similar situation was found in the other Firmicute bacteria which include other important organisms such as the Staphylococciand Clostridia. We provide evidence that the octanoyltransferase of B. subtilis is one of the proteins previously annotated as a lipoate protein ligase and thus this protein, LipM, defines a new class of octanoyltransferases.

EXPERIMENTAL PROCEDURES

Media and Chemicals

Escherichia coli K-12 strains were grown on LB or M9 minimal media (16). Antibiotics were used at the following concentrations (in μg/ml): sodium ampicillin, 100; kanamycin sulfate, 50; chloramphenicol, 12; gentamicin sulfate, 25; tetracycline hydrochloride, 12. All chemicals were obtained from Sigma unless otherwise indicated. Difco Vitamin-Assay Casamino Acids was obtained from Becton-Dickenson. [1-14C]Octanoic acid and [1-14C]octanoyl-CoA were purchased from Moravek. American Radiolabeled Chemicals provided n-[2, 2', 3, 3'-3H]octanoic acid.

Bacterial Strains and Plasmids Constructed

E. coli strains, plasmids and primers used are given in Tables 1 and and2.2. Bacillus subtilis strain 168 was from the Bacillus Genetic Stock Center. Bacterial cultures were grown in shake flasks at 37°C and growth was measured by absorbance at 600 nm using a Beckman DU600 spectrophotometer unless otherwise indicated. Standard techniques (16) for DNA manipulation and cloning were employed unless otherwise indicated. PCR amplification was performed using either Taq (New England Biolabs) or Pfu (Stratagene) polymerases according to the manufacturers recommendations except with the addition of 5% DMSO to the Pfu reactions. The lipB and lplA derivatives of strain EPI300 (Epicentre) were constructed by P1 phage transductions of the mutant alleles from strains QC144 and ZX221 to give strains QC067 and QC068. A functional recA gene was provided by the temperature-sensitive plasmid pEK2. FLP recombinase mediated removal of the antibiotic resistance cassettes was carried out using the temperature-sensitive plasmid pCP20 to yield strain QC069. To obtain a strain deficient in unsaturated fatty acid biosynthesis, a previously described (17) null mutant-fusion derivative of E. coli fabA tagged with chloramphenicol resistance was transduced from strain MH120 to strain W3110 to yield strain QC134. The strain was maintained in LB medium supplemented with 0.5 mM sodium oleate and 0.1% Tergitol NP-40. The fabA allele was also transduced into strain TM131 to yield strain QC168. This strain was maintained on LB medium adjusted to pH 7.0 after supplementation with 5 mM acetate, 5 mM succinate, 0.5 mM oleate and 0.1% Tergitol NP-40.

Table 1
Strains, Plasmids, and Primers Used.
Table 2
Oligonucleotide primers used.

For complementation analysis candidate genes were amplified from genomic DNA by PCR and inserted into pBAD322G (18). The B. subtilis 168 lipM (yqhM) coding sequence was amplified using primers Q005 and Q006 and inserted into the EcoRI and XbaI sites of pBAD322G to give pQC004. E. coli MG1655 lplA was amplified with primers Q017 and Q018 and inserted into the EcoRI and XbaI sites of pBAD322G to give pQC007. The E. coli MG1655 lipB coding sequence was obtained by NcoI and HindIII digestion of pSJ112 (19). The lipB fragment was then inserted into the same sites of pBAD322G to give pQC008. These manipulations placed each of these genes under the control of the arabinose inducible araBAD promoter.

For purification of the lipM protein product the gene was amplified with primers Q022 and Q023 which added a sequence encoding an N-terminal hexahistidine tag. The PCR product was cloned into pET101 using the TOPO cloning kit (Invitrogen). Point mutations within lipM were made using the QuikChange II site directed mutagenesis kit (Stratagene) with primers Q133 to Q140. The gene encoding the B. subtilis 168 glycine cleavage H protein (gcvH or yusH) was PCR amplified with primers Q047 and Q048 which added a C-terminal hexahistidine tag to the gene product. This PCR product was inserted into pCR2.1 using the TA cloning kit (Invitrogen). The gcvH insert was subsequently inserted into the NdeI and SacI sites of pET30a+ to give pQC057. In each of these final constructs the genes were placed under the control of a promoter dependent on phage T7 polymerase (20).

Cosmid Library Selection for Octanoyltransferases

Selection for a complementing cosmid was carried out using the CopyControl Cosmid Library system (Epicentre) as described. Briefly, genomic DNA from B. subtilis 168 was mechanically sheared by repeated pipetting and fragments of about 40 kb were extracted from an agarose gel run overnight at 15 V and gel purified. Size selected DNA was then end repaired and ligated into predigested pCC1FOS vector. The ligation mix was packaged in vitro into λ phage particles and the particles were used to infect strain QC069. After washing three times the cells were titered on LB containing 0.4% glucose, 5 mM acetate, 5 mM succinate, and 24 μg/ml chloramphenicol. To select for lipoic acid prototrophy cells were plated on M9 minimal medium containing 0.4% glycerol, 12 μg/ml chloramphenicol and 0.1 mM each of isoleucine, leucine and valine. The plates were incubated at 37°C for four days and eight colonies were obtained from ca. 580 clones tested. These colonies were reisolated on the same medium to verify their phenotypes and their cosmid were isolated and transformed into strain QC069 to confirm their complementation abilities. The end sequences of the B. subtilis chromosomal fragments were determined by sequencing into the inserts from the flanking vector sequences by the Keck Biotechnology Center using primers purchased from Epicentre.

The growth phenotype of strain QC069 retransformed with the COS1 cosmid was measured. Strains were grown on M9 minimal agar plates of the same composition as that given above and then subcultured overnight in liquid medium of the same composition. The cells were washed thrice with media lacking acetate and succinate then diluted to an OD600 of 0.05. Growth of the cultures (0.4 ml per well) was followed by absorbance at 600 nm using a Bioscreen C instrument with continuous and robust shaking. Growth was measured every 15 min.

Complementation of E. coli lipoate auxotrophs

Genes were expressed in E. coli from plasmids with an arabinose inducible promoter. The strains were first grown on M9 minimal agar and then overnight in liquid with 0.2% arabinose, 0.1% Vitamin-Assay Casamino Acids, 5 mM acetate, 5 mM succinate and gentamicin. Complementation of the lipB lplA strain TM136 was tested using M9 minimal medium containing 0.2% arabinose and gentamicin. Complementation of the lipA lplA strain TM131 was tested using M9 minimal medium containing 0.2% arabinose, gentamicin and 5 μg/ml sodium lipoate. Overnight cultures were washed thrice before subculture to an OD600 of 0.1. Growth curves were obtained using the Bioscreen C instrument as described above for cosmid complementation.

Phylogenetic Analysis and Bioinformatics

Comparison of genes homologous to characterized lipoic acid metabolic genes of different bacteria was done using the SEED database subsystems tool (21). SEED was also used to compare genome context in different organisms and to predict functional coupling.

The phylogeny of the LipB_LplA_BirA family (PF03099) was determined with sequences selected from the Pfam database (22). An unweighted alignment of protein sequences was performed using T-Coffee (23) at the European Bioinformatics Institute website (www.ebi.ac.uk) using the default settings. The edges of the alignment were trimmed using Jalview (24) so only the catalytic domain remained. A minimum evolution tree was constructed with bootstrap analysis of 1000 replicates using Mega4 (25) with the default settings.

Purification of Bacillus subtilis AcpP

The acyl carrier protein of fatty acid biosynthesis was purified from strain EMM99 by a modification of the method previously described (26). All protein steps were carried out at 4°C unless otherwise indicated. Following nickel affinity chromatography the hexahistidine tag was cleaved from approximately 30 mg of ACP with 500 units of AcTEV protease (Invitrogen). The reaction was carried out for 24 h at room temperature during dialysis against the reaction buffer and then subjected to subtractive nickel affinity chromatography as described (26). The modification state of AcpP was verified using conformationally sensitive native PAGE using 20% acrylamide, 2.5 mM urea gels buffered with Tris-glycine (27). The TEV protease-digested AcpP migrated much slower than the undigested tagged protein. Fractions containing the TEV-cleaved apo and holo AcpP were pooled, concentrated with a Vivaspin concentrator (GE Healthcare) and flash frozen for storage at -80°C. Quantitative conversion of the mixture to holo-ACP was obtained using the B. subtilis Sfp phosphopantetheinyl transferase as previously described (28). Strain QC120 was grown in LB with kanamycin and induced for 2 h with 1 mM isopropyl β-d-1-thiogalactopyranoside at an OD600 of 0.6. Sfp was purified from strain QC120 extracts by nickel affinity chromatography as described for LplA (29). Pooled fractions containing Sfp were precipitated with ammonium sulfate (35% to 55% of saturation) as described (28) and dialyzed overnight against 50 mM sodium phosphate (pH 7.0), 10 mM MgCl2, 10% glycerol and 1 mM DTT. The protein was concentrated with a Vivaspin concentrator (GE Healthcare) and flash frozen for storage at --80°C. Conversion to holo-ACP was verified using conformationally sensitive gel electrophoresis as before (27). The AcpP modification state was also assayed by electrospray ionization mass spectrometry performed by the University of Illinois School of Chemical Sciences Mass Spectrometry Facility. The sample was prepared by dialysis overnight against 10 mM ammonium bicarbonate and then the solvent was removed by evaporation under a stream of nitrogen. The mass was determined by electrospray ionization mass spectrometry in positive ion mode using a Micromass Quattro II with 100 pmol/ml of protein in 50% aqueous acetonitrile containing 0.1% formic acid.

Preparation of 3H-Octanoyl-ACP

Octanoyl-ACP was prepared using Vibrio harveyi acyl ACP synthetase (AasS) purified by nickel affinity and ion exchange chromatography as previously described (30). The 100 μl reaction mixture contained 10 mM Tris-HCl (pH 8.0), 10.1 mM MgCl2, 10 mM ATP, 1 mM TCEP, 5 mM sodium [3H]octanoate, 0.5 mM holo-ACP, and 5 μM AasS. The reaction was allowed to proceed for two h at 37°C. The product was precipitated at 4°C overnight after addition of an equal volume of acetone. The pellet was resuspended in 100 mM Tris-HCl (pH 8.0) and dialyzed overnight against 10 mM sodium 2-(N-morpholino)ethanesulfonic acid (pH 6.1). The protein was found to be >95% octanoyl-ACP by gel electrophoresis using conformationally sensitive native PAGE (27). Proteins were quantified by use of extinction coefficients calculated using the program PROTPARAM on the EXPASY website (31). Octanoyl-ACP was quantified at 280 nm using the calculated extinction coefficient of 1,490 M-1 cm-1.

Purification of B. subtilis GcvH

To purify hexahistidine-tagged B. subtilis GcvH the lipoic acid auxotrophic strain QC146 was used. The strain was grown in LB with 0.1% glucose, 5 mM acetate, and 5 mM succinate (pH 7.0) to an absorbance of 0.5 at 600 nm. Expression was induced with 0.2% arabinose and the culture incubated for another 4 h before the cells were pelleted by centrifugation and frozen at -80 °C. The protein was purified by nickel affinity and anion exchange chromatographic steps and analyzed by mass spectroscopy as described above for AcpP. GcvH was quantified at 280 nm using the calculated extinction coefficient of 16,960 M-1 cm-1.

Purification of LipM

LipM was initially purified from strain QC103 by nickel affinity and anion exchange chromatographic steps as previously described for T. acidophilum LplA (29) except that the reducing agent was 5 mM TCEP instead of DTT. Unmodified LipM was obtained using the fabA strain QC142. The strain was grown in LB with 0.1% glucose, 0.5 mM sodium oleate, 0.1% Tergitol NP-40, ampicillin and kanamycin to an OD600 of 0.5. At this point T7 polymerase expression was induced by addition of 0.2% arabinose for 1.5 h. Then fatty acid biosynthesis was inhibited by addition of 0.1 μg/ml triclosan for 40 min. Cells were harvested by centrifugation and frozen at -80°C. Lysis and purification were performed as for the initial purification. An calculated extinction coefficient of 45,380 M-1 cm-1 at 280 nm was used for LipM. The purified protein was analyzed by matrix-assisted laser desorption/ionization (MALDI) mass spectrometry at the University of Illinois School of Chemical Sciences Mass Spectrometry Facility. The sample was prepared by dialysis overnight in 10 mM ammonium acetate, dried under a stream of nitrogen and subjected to MALDI performed using a Voyager-DE STR mass spectrometer (Applied Biosystems) equipped with a UV laser (337 nm N2 laser). All measurements were made using the linear mode and the positive ion was recorded. Samples were prepared by mixing 2 μl of sample and 10 μl of the matrix, α-cyano-4-hydroxycinnamic acid.

Liquid Chromatography Tandem Mass Spectrometry Analysis of LipM

LipM was purified from strain QC103 as described for LplA (29) and was subjected to size exclusion chromatography using a Sephadex 200 column developed with 10 mM sodium phosphate (pH 7), containing 100 mM sodium chloride and 1 mM DTT. Fractions containing pure LipM as judged by SDS PAGE were pooled, dialyzed against 10 mM ammonium acetate and dried under a stream of nitrogen. The protein was then dissolved in 100 mM Tris-HCl (pH 8.5), containing 1 M urea and 20 mM methylamine and digested with one-tenth mass of Modified Sequencing Grade Trypsin (Roche) for 16 h at 37°C. Liquid chromatography tandem mass spectrometric analysis was performed by the University of Illinois Biotechnology Center Protein Sciences Facility on a Waters Q-Tof API-US Quad-ToF mass spectrometer with a nanoAcquity UPLC system. The columns used were Waters nanoAcquity UPLC (75 micron x 150 mm 3 μm Atlantis dC18) and Atlantis dC18 5 μm Nanoease trap columns. A 60 min linear gradient of 1% to 60% acetonitrile in 0.1% formic acid was used to elute the peptides from the columns. Tandem mass spectrometric data were collected using the Data Directed Analysis method in MassLynx to fragment the top four ions in each survey scan. ProteinLynx (Waters) was used to process the mass spectral data into peak list files for analysis by Mascot (Matrix Science). Database searches were performed against the NCBI non-redundant database.

Assay of Enzyme Bound Fatty Acids

LipM was purified from two liter cultures with and without addition of triclosan by nickel affinity chromatography as described above. The eluates were twice dialyzed against 10 mM ammonium acetate first overnight and again for 4 h. The dialysate was evaporated under a stream of nitrogen with heating to 42°C. A modification of previously described syntheses of fatty acid butyl esters was then carried out (11, 32). To each glass vial was added 500 μl hexanes, 500 μl butanol-BF3 and 100 μg anhydrous magnesium sulfate. Heptanoic acid (10 μg) was added as an internal standard. Transesterification was carried out at 65°C for 2 h. Salts and butanol were removed by three extractions with water. Butyl esters were analyzed by gas chromatography mass spectrometry (GC-MS) by a modification of the method of Hermes and Cronan (11).

Samples (5 μL) were injected in split-less mode to the GC/MS system consisted of an Agilent 7890A gas chromatograph, an Agilent 5975 mass selective detector and Agilent 7683B (Agilent Inc, Palo Alto, CA, USA) autosampler. Injections were performed on a 30 m ZB-WAX column with 0.32 mm I.D. and 0.25 mm film thickness (Phenomenex) with an injection port temperature of 230°C, the interface set to 250°C, and the ion source adjusted to 230°C. The helium carrier gas was set at a constant flow rate of 3 ml min-1. The temperature program was 5 min isothermal heating at 90°C followed by an oven temperature increase of 5 °C min-1 to 260 °C for 10 min. The mass spectrometer was operated in positive electron impact mode (EI) at 69.9 eV ionization energy in m/z 30-800 scan range. The spectra of all chromatogram peaks evaluated using the HP Chemstation (Agilent) and AMDIS (NIST, Gaithersburg, MD, USA) programs. The spectra of all chromatogram peaks were compared with electron impact mass spectrum libraries NIST08 (NIST, MD, USA) and WILEY08 (Palisade Corp.).

To determine the efficiency of esterification octanoyl-LipM was made in vitro. AasS was used to generate octanoyl-ACP in the LipM reaction mixture. The 200 μl reactions contained 100 mM sodium phosphate (pH 7.0), 50 mM NaCl, 5 mM TCEP, 10 mM ATP, 10 mM MgCl2, 10 mM [14C]octanoate, 50 μM holo-ACP, 5 μM AasS, and 5 μM LipM. After incubation at 37°C for 2 h the reaction and the control reaction lacking LipM were dialyzed. The proteins were dried, esterified and analyzed by GC-MS as described above.

Assay of LipM catalyzed Octanoyltransfer

Transfer of octanoate from octanoyl-ACP to GcvH was assayed using a coupled system in which AasS provided the octanoyl-ACP substrate. The 25 μl reactions contained 100 mM sodium phosphate (pH 7.0), 50 mM NaCl, 5 mM TCEP, 1 mM ATP, 1 mM MgCl2, 250 μM [14C]octanoate, 50 μM holo-ACP, 20 μM GcvH, 2 μM AasS, and 1 μM LipM. When formation of acyl-enzyme intermediate was studied, the concentrations of wild type and mutant LipM proteins were increased to 10 μM. To test octanoyl-CoA as a LipM substrate a coupled assay was also used. The 25 μl reactions contained 100 mM sodium phosphate (pH 7.0), 50 mM NaCl, 5 mM TCEP, 1 mM ATP, 1 mM MgCl2, 1 mM [14C]octanoate, 50 μM trilithium CoA, 20 μM GcvH, 0.1 unit acyl-CoA synthetase (Pseudomonas sp. from Sigma) and 1 μM LipM. The reactions were incubated at 37°C for one h and subjected to SDS-PAGE using a loading buffer that lacked reducing agents and 4-20% Trisglycine gels. The gels were stained with Coomassie R-250, soaked in Amplify fluorographic reagent (GE Healthcare), and dried onto Whatman filter paper. Dried gels were exposed to Biomax XAR film (Kodak) flashed with an Amersham Sensitize Preflash Unit (GE Healthcare) (33). Exposure was carried out for 16-24 h at -80°C.

Isolation and transfer of the octanoyl moiety of the octanoyl-LipM intermediate

Synthesis of the octanoyl-LipM intermediate was done in a 500 μl assay containing 50 mM sodium phosphate (pH 7.0), 5 mM TCEP, 10 mM MgCl2, 10 mM ATP, 1 mM [14C]octanoate, 0.1 mM holo-ACP, 2 μM AasS, and 10 μM LipM. The reaction was incubated at 37°C for one h. The product was diluted 10-fold in 25 mM Tris-HCl (pH 8.0) and purified by anion exchange chromatography as described for LplA (29). LipM eluted at about 110 mM sodium chloride, the same point as apo-LipM. Fractions containing pure LipM were pooled, concentrated and exchanged into 10 mM sodium phosphate (pH 7.0) using a Vivaspin concentrator (GE Healthcare). The transfer reaction from octanoyl-LipM to GcvH or holo-ACP contained 100 mM sodium phosphate (pH 7.0), 5 mM TCEP, 15 μM LipM and either GcvH or holo-ACP at 20 μM. The reactions were incubated at 37°C for one h and visualized by SDS-PAGE and radiography as previously described for the LipM assay.

Competition assay using CoA and ACP octanoyl thioesters

LipM was assayed for the ability to utilize a mixture of octanoyl-ACP and octanoyl-CoA in a competition assay similar to that described previously (11). The 100 μl reactions contained 10 mM sodium phosphate (pH 7.0), 100 mM sodium chloride, 5 mM TCEP, 250 μM [14C]octanoyl-CoA, 50 μM 3H-octanoyl-ACP, 100 μM GcvH, 2 μM AasS, and 1 μM LipM. The reaction mixture was preincubated at 37°C for 3 min before addition of LipM. After 30 min at 37°C the assay was stopped by addition of guanidine HCl to 5 M. The entire reaction was applied to a Ni-NTA spin column (Qiagen) prequilibriated with 50 mM Tris-HCl (pH 8.0) containing 4 M urea. The column was washed four times with 600 μl of the same buffer. Hexahistidine tagged LipM and GcvH were eluted with 400 μl of the same buffer containing 500 mM imidazole. The eluate was diluted to 1 ml with water and mixed with 4 ml of Biosafe II scintillation cocktail. The mixture was subject to scintillation counting using a Beckman LS 6500. The 3H and 14C isotopes were differentially counted using channels 0-400 and 0-670 respectively. The 3H counts were adjusted by subtracting 67% of the 14C counts for that sample, which was experimentally determined from six different amounts of 14C octanoyl-CoA in the assay elution buffer.

RESULTS

Genetic complementation of an E. coli lipB strain

Although amino acid homology is a useful tool to predict function, the B. subtilis genome encoded no recognizable homologue of LipB. Therefore, in order to find the gene(s) responsible for the octanoyltransfer step of the B. subtilis lipoic acid synthetic pathway we constructed a library of B. subtilis chromosomal fragments in a cosmid vector and used the resulting phage particles to transfect an E. coli lipB lplA double mutant strain. The lplA mutation was introduced into this strain in order to avoid possible bypass of the lipB mutation by LplA-catalyzed ligation of traces of intracellular octanoate or lipoate (11). Glycerol was used as the carbon source to preclude bypass of succinate plus acetate-dependent growth by fermentative metabolism. The B. subtilis genomic library contained a sufficient number of clones to cover 99.6% of the genome as calculated from the equation of Clarke and Carbon (34). The eight cosmid clones that allowed growth of the lipB lplA strain carried overlapping fragments of the B. subtilis genome (Table 3) that contained a common region in which only a single candidate gene related to lipoic acid metabolism was found; the yqhM gene annotated as encoding a lipoate ligase. To test if yqhM was the gene responsible for complementation by the cosmid clones, the minimal gene was expressed from an arabinose inducible promoter. Expression of yqhM (or E. coli lipB or E. coli lplA) allowed growth of lipB lplA double mutant strain, TM136 (Figure 2). Since the growth observed could either be due to octanoyltransferase activity or to high-level lipoyl ligase activity, we also expressed lipM in strain TM131, a lipA lplA strain, in the presence of lipoic acid. Expression of yqhM (or E. coli lipB) failed to allow growth of this strain whereas expression of E. coli lplA gene resulted in robust growth. Therefore, despite its sequence similarity to E. coli LplA (Figure 8, and and9),9), YqhM lacks lipoate ligase activity and instead has octanoyltransferase activity (Figure 2). Therefore we have renamed yqhM as lipM.

Figure 2
Complementation of E. coli Lipoic Acid Auxotrophs. Growth curves of lipoic acid auxotrophs carrying various plasmids are shown. Panel A. Growth tests for lipB complementation in minimal glycerol medium. Growth curves of the wild type (WT) and lipB lplA ...
Figure 8
Alignment of LipM with Enzymatically Characterized LipBs and LplA Proteins. Alignments were performed as described in Experimental Procedures and displayed using Jalview (24) . The sequence name indicates the enzyme type, the Uniprot code indicates the ...
Figure 9
Phylogeny of LipM. The minimum evolution tree of selected PF03099 protein sequences with bootstrap percentage confidence values shown for each branch is given. Phylogenetic analyses were done as described in Experimental Procedures. The scale bar corresponds ...
Table 3
B. subtilis 168 genome segments in the complementing cosmids.

Purification of GcvH

To provide a substrate for in vitro studies of LipM the putative glycine cleavage H protein (GcvH) of Bacillus subtilis 168 was purified in its apo form from an E. coli strain deficient in lipoic acid biosynthesis. The pure protein formed a doublet when analyzed by SDS-PAGE (Figure 3A), but electrospray mass spectrometry gave a major peak at 14,944.2 amu (Figure 3B). This is 1.2 amu less than that theoretically predicted which we attribute to deamidation of the protein (which results in a negligible mass difference) perhaps at N73 which since it is followed by a glycine residue should be unusually labile (35). Deamidation of other LDs has been observed and does not affect modification (36). Note that GcvH migrates aberrantly in SDS-PAGE analysis (Figure 3A). It runs as though it is twice its actual size of 14.1 kDa. Such slow migration rates are often seen with small, very acidic proteins (GcvH has a calculated pI of 3.9) and can be attributed to abnormally low SDS binding.

Figure 3
Analysis of Purified Proteins. Panel A: SDS PAGE of 0.2 μmol each of purified LipM and GcvH. Molecular weight standards are indicated in kDa. Panel B: electrospray ionization mass spectrum of GcvH. Calculated masses are represented by black circles. ...

Modification of LipM and Purification of Apo Protein

To directly test LipM for octanoyltransferase activity we purified the protein obtained after high-level expression in E. coli and it appeared homogenous by SDS-PAGE (Figure 3A) and size exclusion chromatography where it eluted as a protein of 14 kDa mainly in the monomeric form with a variable small secondary peak that may be a multimer (Figure 3D). However, the protein obtained in the initial purifications of LipM was inactive and had an average molecular weight of 33,338 +/-32 (5 measurements), a value 143 amu greater than expected which suggested that the protein had been post-translationally modified (the initiator methionine was retained as expected from the specificity of E. coli methionine aminopeptidase (37, 38)). However, LipM gave rather imprecise MALDI mass spectra which may have resulted from a combination of a heterogeneous set of modifications plus the smoothed and averaged data generated by the spectrometer (Figure 3C). In order to determine if the protein was postranslationally modified, we digested LipM with trypsin and subjected the resulting peptides to liquid chromatography tandem mass spectrometry analysis. We found evidence for a series of modified peptides that had overlapping sequences and all contained residue C150. In some LipM molecules the predicted C150 modification was an octanoyl thioester whereas in others a decanoyl adduct was present (Figure 4). By analogy to LipB these data suggested that C150 is the LipM active site nucleophile and the octanoyl thioester modification is the acyl-enzyme intermediate (4).

Figure 4
Liquid Chromatography Tandem Mass Spectrometric Analysis of LipM tryptic peptides. Panel A. Modification states of C150 detected. The theoretical peptides are shown with C150 in bold and underline type. The difference in mass from the modification is ...

The decanoyl adduct was previously found as a modification of Mycobacterium tuberculosis LipB when the protein was expressed in E. coli, but not when expressed in a mycobacterium (13). In that work it was proposed (albeit, not proven) that the M. tuberculosis LipB decanoic acid adduct arose by binding cis-3-decenoyl-ACP, a key intermediate in E. coli fatty acid biosynthesis by LipB followed by Michael addition of the active site cysteine thiol to the double bond resulting in a thioether link to C-3 of the acyl chain. Since like M. tuberculosis, the B. subtilis unsaturated fatty acid synthetic pathway differs from that of E. coli, it seemed likely that in both cases the octanoyltransferase would accidently bind cis-3-decenoyl-ACP and the decanoyl adduct would form. To avoid decanoyl adduct formation we expressed LipM in an E. coli fabA mutant strain. This strain lacked the ability to make cis-3-decenoyl-ACP (hence unsaturated fatty acids) and therefore growth of the strain required supplementation of the medium with an unsaturated fatty acid such as oleic acid (17).

To reduce the level of the octanoyl-enzyme intermediate we added triclosan, an inhibitor of fatty acid biosynthesis (39) to the medium during LipM expression. We determined the levels of LipM with bound octanoate by GC-MS of butyl esters using an octanoyl-LipM standard prepared in vitro. Using this standard a recovery efficiency of 93% based on complete conversion of LipM to octanoyl-LipM was found, which is within the expected efficiency for esterification (32). This method showed that triclosan addition resulted in a significant reduction in the level of octanoyl-enzyme (Figure 5) and thus by triclosan addition and use of the fabA strain we obtained LipM in a largely unmodified and active form.

Figure 5
Gas Chromatography Mass Spectrometric Analysis of LipM Bound Octanoate. LipM-bound octanoyl moieties were assayed by transesterification to the butyl esters followed by GC-MS analysis as described in Experimental Procedures. Panel A: The values are the ...

In vitro demonstration of LipM activity

We employed the AasS acyl-ACP synthetase to generate acyl-ACPs in the LipM assay mixture and found that octanoyl transfer from octanoyl-ACP to B. subtilis GcvH occurred in a LipM dependent manner (Figure 6). In the absence of the GcvH acceptor protein, LipB was converted to the octanoyl-enzyme intermediate. The octanoyl-LipM species was purified and shown to transfer the octanoyl moiety to either GcvH in the forward reaction or to ACP is the reverse reaction, as expected of a catalytically competent intermediate. Due to the lengthy time of incubation we expect that the relative levels of transfer in the forward and reverse directions reflect the equilibria of the transfer reactions. We also tested octanoyl-CoA as an octanoyl donor by using acyl-CoA synthetase to generate the substrate in the reaction mix. This system was used to demonstrate that LipM also uses octanoyl-CoA as a substrate, albeit poorly (Figure 7). This is not unexpected as CoA is a substrate mimic of ACP. In order to determine which of the substrates LipM preferred we performed a double label experiment with equal concentrations of octanoyl-CoA and octanoyl-ACP differentially labeled in the octanoyl moiety. Octanoyl-ACP was much the preferred substrate for modification of GcvH, although some modification by transfer from octanoyl-CoA was observed (Figure 7).

Figure 6
Octanoyltransferase Activities of Wild Type and Mutant LipM Proteins. Assay of octanoyl transfer from octanoyl-ACP to B. subtilis GcvH using either [1-14C]octanoyl-ACP or octanoyl-ACP as substrates as described in Experimental Procedures. Autoradiograms ...
Figure 7
Comparison of the octanoyl thioesters of ACP and CoA as LipM Substrates. Assays of octanoyltransfer to B. subtilis GcvH using either octanoyl-CoA or octanoyl-ACP as described in Experimental Procedures. Panel A: Autoradiograms of dried SDS-PAGE gels are ...

C150 and K165 are key LipM catalytic residues

We constructed point mutations in lipM in order to test the importance of the predicted active site residues. The residues mutated were C150 and the conserved lysine, K165 in LipM, which is present in all members of PF03099 (14). Mutations C150A or C150S resulted in loss of the overall catalytic activity and the inability to form the acyl-enzyme intermediate whereas K165A had reduced catalytic ability and K165R remained active under these conditions (Figure 6C).

Bioinformatic analysis of LipM

Phylogenetic analysis of LipM and proteins of a high degree of sequence similarity revealed that these proteins form a clade distinct from other functional groups of related enzymes, as well as being distinct from LipB type octanoyltransferases (Figure 9). This suggests that other members of the LipM clade are also octanoyltransferases. The LipM clade groups with lipoate protein ligases with 73% bootstrap support, which is considered significant (40). This means that LipMs are closely related to LplAs and likely share a more recent ancestor with LplAs than with any other group of proteins consistent with the current incorrect annotation of LipM as a lipoate protein ligase.

Close homologues of LipM are found in Firmicutes and Cyanobacteria (Figure 9). Although relative to LipB the cyanobacterial LipMs form a separate clade from Firmicute LipMs, they have a similar insertion that contains the active site cysteine as in B. subtilis LipM. This and the high amino acid sequence similarity (~50%) suggest that the cyanobacterial LipMs are octanoyltransferases. The Cyanobacteria also encode a putative LipB type octanoyltransferase which (if both enzymes are active) would give these bacteria a pair of octanoyltransferase isozymes. The presence of two isozymes in Cyanobacteria suggests that there may be a physiological difference between the two enzymes within at least these organisms. Further work is necessary to determine if there are physiologically relevant differences between the LipB and LipM types of octanoyltransferase.

Using the SEED subsystem database (21) we found that Firmicutes and Cyanobacteria also encode putative lipoyl synthases and thus the sulfur insertion enzyme seems present. Moreover the SEED database shows that putative Firmicute lipM genes are functionally coupled to the glycine cleavage system; lipM is very often found adjacent to genes encoding putative glycine cleavage system P and T proteins. The covariance with putative LipA homologues and the functional coupling with the glycine cleavage system provide further indications that these LipM homologues are involved in lipoic acid biosynthesis.

DISCUSSION

Using an approach unbiased by bioinformatics we have identified a B. subtilis gene encoding a new type of octanoyltransferase. Our isolation of lipM on eight different cosmid clones suggests that it is the only B. subtilis 168 gene able to complement E. coli lipB, although this suggestion must be tested by genetic analyses. A B. subtilis strain lacking LipM activity should be a lipoate auxotroph assuming that the yhfJ gene encodes a lipoate ligase (Figure 1) (11). We and others have found that overproduction of E. coli lplA complements lipB null mutants (Figure 2) (7, 11) and thus we would expect to have also isolated the yhfJ gene. However, this was not the case. The most straightforward explanation for this result was that the level of yhfJ expression was insufficient to restore growth under our experimental conditions. Either high-level expression of wild type lplA or normal expression of an LplA mutant protein having increased octanoate affinity is required for lipB complementation (Figure 2) (11). Other possibilities are that our library is biased and lacked yhfJ clones or that yhfJ does not encode a lipoate ligase. We favor the biased library explanation because this would be consistent with the unexpectedly frequent isolation of LipM clones. Library bias due to gene toxicity is a known problem with B. subtilis genomic DNA libraries in E. coli was even seen with a chromosomally integrated library (41). Although we used a vector with a single copy F factor replication origin vector, basal expression from the second trfA-dependent replication origin (42) may also have been an issue. Indeed, growth of most isolates of our cosmid library was much better on glycerol or glucose minimal media than on medium containing arabinose (data not shown) which would induce trfA expression resulting in greatly increased plasmid copy numbers (42).

Upon purification of LipM we found that the average mass determined by MALDI was greater than that calculated from the amino acid sequence and this, plus the precedent of the M. tuberculosis LipB decanoic acid adduct (13), strongly suggested that the protein was modified. Analysis of the LipM tryptic peptides indicated that both the decanoic acid adduct and the octanoyl-LipM intermediate were present in these preparations. The level of octanoyl-enzyme intermediate was significantly decreased by addition of the fatty acid synthesis inhibitor, triclosan (which inhibits FabI, the enzyme responsible for the last step of the fatty acid synthesis cycle) during LipM expression. The uncertainties of peptide ionization efficiencies precluded quantitation of the levels of octanoylation by analysis of tryptic peptides and thus we developed a method to directly detect the octanoyl groups as their butyl esters, a derivative that allows efficient recovery of short chain volatile acids such as octanoate (11). To our knowledge this is the first time this method has been adapted for quantification of a protein modification.

Despite their marked differences in sequence the LipM and LipB octanoyltransferases share the same general catalytic mechanism. The octanoyl moiety is transferred from octanoyl-ACP to the enzyme active site cysteine thiol to give octanoyl-enzyme the thioester bond of which is attacked by the ε-amino group of the LD lysine residue to give octanoyl-LD. The octanoyl moiety can also be transferred from the acyl enzyme intermediate back to ACP. However, the results of mutagenesis studies distinguish the detailed LipM mechanism from that of LipB. Although loss of the active site thiol by substitution of the cysteine residue with alanine or serine results in inactivation of both enzymes, the E. coli LipB C169A protein performs dead-end acylation of the enzyme (4) whereas LipM C150A does not. Moreover, the LipB C169S protein forms a dead end octanoyl ester (4) whereas LipM C150S does not. This may simply be a result of the different environments of the active site nucleophile or it may reflect a physiologically relevant mechanism to impart resistance against self-inactivation to LipM. The members of the PF03099 enzyme family have a strictly conserved lysine residue the only totally conserved residue of the protein family (14). Loss of the charge of the conserved lysine side chain inactivates M. tuberculosis LipB (13) whereas the LipM mutant protein retained some activity (Figure 6). From the available structures of LplA and LipB proteins it appears that this residue is near the reactive carbonyl of the substrate. This suggests it is important for stabilization of the oxyanion, as is the case in the mononucleotide binding fold (43). Other positively charged lysine and arginine side chains are in the vicinity of K165, which may explain why the LipM K165A mutant retains trace activity. The activity of the LipM K165R protein supports this notion because substitution of the longer positively charged side chain results in a protein with activity similar to that of the wild type protein.

It is interesting that LipM and LipB, together with the LplA-LplB bipartite lipoate ligase (11), share the property of anomalous migration in size exclusion chromatography. The three proteins all migrate as though they are about half of their known sizes. LipM elutes as a protein of 14 kDa whereas E. coli LipB elutes as though it were a protein of 10 kDa (29, 44). The most straightforward explanation of this unusual behavior is that these proteins interact with the chromatographic matrix within the bead and thus emerge from the matrix more slowly than expected (this behavior is independent of the salt concentration of the eluate (44)). Given these data it seems likely that both LipM and LipB are monomeric proteins, although confirmatory data obtained by another method (e.g., analytical ultracentrifugation) are clearly needed.

As discussed above, LipB octanoyltransferases and LplA lipoate protein ligases have similar structures and together with the biotin protein ligases make up the divergent enzyme family PF03099. The LipM type of octanoyltransferase provides a bridge between the LipB and LplA proteins, in that LipM has the activity of LipB, but an amino acid sequence that resembles LplA. Although LipM has sequence homology with E. coli LplA, it is a considerably shorter protein. The characterized LipB octanoyltransferases are composed of a single domain whereas canonical lipoate protein ligases possess a catalytic domain of a size similar to that of LipB plus a C-terminal accessory domain. Since nearly half of the sequences in the LplA subfamily contain only a single catalytic domain (29), it is tempting to designate all of these “short LplAs” as octanoyltransferases. However, the accessory domain can also be a separate protein as recently demonstrated for the Thermoplasma acidophilum lipoate ligase (29, 45). Although the absence of an accessory domain indicates that the short proteins annotated as lipoate ligases lack lipoyl ligase activity (29), for any given protein remains possible that the protein is inactive due to loss of its accessory domain protein. In the known case of a bipartite lipoate ligase, the accessory protein is encoded by a gene adjacent to the LplA homologue (29). However, there is no reason that this must be the case (although it could simplify stoichiometric expression of the subunits). Indeed, there are putative bipartite ligases encoded by genes located far apart in the genome, such as in Pyrococcus furiosus (29).

The distinct phylogeny of LipM proteins should allow more correct prediction and annotation of genes currently annotated as encoding short lipoyl ligases. Since the octanoyltransferases and lipoate ligases perform different chemistry, octanoyltransferases and ligases could be distinguished by the octanoyltransferase active site cysteine and the neighboring residues. However, the residues that flank the active site cysteine residues of LipM and LipB show no similarity. Indeed, relative to LipB the C150 region of LipM seems to reside on an inserted loop (Figure 8). As such, they are different octanoyltransferase types and must be considered separately.

The LipM and LipB octanoyltransferases can be added to the list of enzymes that have undergone convergent evolution functionally and mechanistically (46, 47). As previously stated, these enzymes come from the same family and have a similar protein fold. Despite this, they have distinct primary structures indicating divergence at the sequence level. Therefore, it appears that octanoyltransferase activity has evolved from the same protein scaffold by two different paths. This highlights the multitude of evolutionary outcomes that are possible which explains the rarity of true convergent evolution at the sequence level (46)

ACKNOWLEDGMENTS

We thank Drs. Nicholas De Lay, Mariano Martinez and Yanfang Jiang for experimental materials.

Abbreviations

LD
lipoyl domain
AasS
acyl-ACP synthetase soluble
ACP
acyl carrier protein
FabA
3-hydroxydecanoyl-ACP dehydratase:trans-2-decenoyl-ACP isomerase
TCEP
(tris(2-carboxyethyl)phosphine)
DTT
dithiothreitol
GcvH
Glycine cleavage system H protein

Footnotes

This investigation was supported by the National Institutes of Health under Ruth L. Kirschstein National Research Service Award 5 T32 GM070421 from the National Institute of General Medical Sciences and grant AI15650 from the National Institute of Allergy and Infectious Disease.

REFERENCES

1. Perham RN. Swinging arms and swinging domains in multifunctional enzymes: catalytic machines for multistep reactions. Annu. Rev. Biochem. 2000;69:961–1004. [PubMed]
2. Xiao Z, Xu P. Acetoin metabolism in bacteria. Crit Rev Microbiol. 2007;33:127–140. [PubMed]
3. Cronan JE, Zhao X, Jiang Y. Function, attachment and synthesis of lipoic acid in Escherichia coli. Advances in Microbial Physiology. 2005:103–146. [PubMed]
4. Zhao X, Miller JR, Cronan JE. The reaction of LipB, the octanoyl-[acyl carrier protein]:protein N-octanoyltransferase of lipoic acid synthesis, proceeds through an acyl-enzyme intermediate. Biochemistry. 2005;44:16737–16746. [PubMed]
5. Douglas P, Kriek M, Bryant P, Roach PL. Lipoyl synthase inserts sulfur atoms into an octanoyl substrate in a stepwise manner. Angew Chem Int Ed Engl. 2006;45:5197–5199. [PubMed]
6. Cicchillo RM, Booker SJ. Mechanistic investigations of lipoic acid biosynthesis in Escherichia coli: Both sulfur atoms in lipoic acid are contributed by the same lipoyl synthase polypeptide. J. Am. Chem. Soc. 2005;127:2860–2861. [PubMed]
7. Morris T, Reed K, Cronan J. Lipoic acid metabolism in Escherichia coli: the lplA and lipB genes define redundant pathways for ligation of lipoyl groups to apoprotein. J. Bacteriol. 1995;177:1–10. [PMC free article] [PubMed]
8. Morris TW, Reed KE, Cronan JE., Jr. Identification of the gene encoding lipoate-protein ligase A of Escherichia coli. Molecular cloning and characterization of the lplA gene and gene product. J Biol Chem. 1994;269:16091–16100. [PubMed]
9. Reed LJ, Leach FR, Koike M. Studies on a lipoic acid-activating system. J. Biol. Chem. 1958;232:123–142. [PubMed]
10. Green D, Morris T, Green J, Cronan J, Jr, Guest J. Purification and properties of the lipoate protein ligase of Escherichia coli. Biochem. J. 1995;309:853–862. [PMC free article] [PubMed]
11. Hermes F, Cronan JE. Scavenging of cytosolic octanoic acid by mutant LplA lipoate ligases allows growth of Escherichia coli strains lacking the LipB octanoyltransferase of lipoic acid synthesis. J. Bacteriol. 2009;191:6796–6803. [PMC free article] [PubMed]
12. Zhao X, Miller JR, Jiang Y, Marletta MA, Cronan JE. Assembly of the covalent linkage between lipoic acid and its cognate enzymes. Chem Biol. 2003;10:1293–1302. [PubMed]
13. Ma Q, Zhao X, Eddine AN, Geerlof A, Li X, Cronan JE, Kaufmann SHE, Wilmanns M. The Mycobacterium tuberculosis LipB enzyme functions as a cysteine/lysine dyad acyltransferase. Proc. Natl. Acad. Sci. USA. 2006;103:8662–8667. [PMC free article] [PubMed]
14. Reche PA. Lipoylating and biotinylating enzymes contain a homologous catalytic module. Protein Sci. 2000;9:1922–1929. [PMC free article] [PubMed]
15. Martin N, Lombardia E, Altabe SG, de Mendoza D, Mansilla MC. A lipA (yutB) mutant, encoding lipoic acid synthase, provides insight into the interplay between branched-chain and unsaturated fatty acid biosynthesis in Bacillus subtilis. J Bacteriol. 2009;191:7447–7455. [PMC free article] [PubMed]
16. Sambrook J, Russell D. Molecular cloning: a laboratory manual. CSHL Press; 2001.
17. Henry MF, Cronan JE. A new mechanism of transcriptional regulation: Release of an activator triggered by small molecule binding. Cell. 1992;70:671–679. [PubMed]
18. Cronan JE. A family of arabinose-inducible Escherichia coli expression vectors having pBR322 copy control. Plasmid. 2006;55:152–157. [PubMed]
19. Jordan SW, Cronan JE., Jr. The Escherichia coli lipB gene encodes lipoyl (octanoyl)-acyl carrier protein:protein transferase. J Bacteriol. 2003;185:1582–1589. [PMC free article] [PubMed]
20. Studier FW, Moffatt BA. Use of bacteriophage T7 RNA polymerase to direct selective high-level expression of cloned genes. J Mol Biol. 1986;189:113–130. [PubMed]
21. Overbeek R, Begley T, Butler RM, Choudhuri JV, Chuang H-Y, Cohoon M, de Crecy-Lagard V, Diaz N, Disz T, Edwards R, Fonstein M, Frank ED, Gerdes S, Glass EM, Goesmann A, Hanson A, Iwata-Reuyl D, Jensen R, Jamshidi N, Krause L, Kubal M, Larsen N, Linke B, McHardy AC, Meyer F, Neuweger H, Olsen G, Olson R, Osterman A, Portnoy V, Pusch GD, Rodionov DA, Ruckert C, Steiner J, Stevens R, Thiele I, Vassieva O, Ye Y, Zagnitko O, Vonstein V. The subsystems approach to genome annotation and its use in the project to annotate 1000 genomes. Nucl. Acids Res. 2005;33:5691–5702. [PMC free article] [PubMed]
22. Finn RD, Mistry J, Tate J, Coggill P, Heger A, Pollington JE, Gavin OL, Gunasekaran P, Ceric G, Forslund K, Holm L, Sonnhammer ELL, Eddy SR, Bateman A. The Pfam protein families database. Nucl. Acids Res. 2010;38:D211–222. [PMC free article] [PubMed]
23. Notredame C, Higgins DG, Heringa J. T-coffee: a novel method for fast and accurate multiple sequence alignment. J. Mol. Biol. 2000;302:205–217. [PubMed]
24. Waterhouse AM, Procter JB, Martin DMA, Clamp M, Barton GJ. Jalview Version 2--a multiple sequence alignment editor and analysis workbench. Bioinformatics. 2009;25:1189–1191. [PMC free article] [PubMed]
25. Tamura K, Dudley J, Nei M, Kumar S. MEGA4: Molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol Biol Evol. 2007;24:1596–1599. [PubMed]
26. Martinez MA, Zaballa M. a.-E., Schaeffer F, Bellinzoni M, Albanesi D, Schujman GE, Vila AJ, Alzari PM, de Mendoza D. A Novel Role of Malonyl-ACP in Lipid Homeostasis. Biochemistry. 2010;49:3161–3167. [PubMed]
27. Cronan JE, Thomas J. Bacterial fatty acid synthesis and its relationships with polyketide synthetic pathways. Methods Enzymol. 2009;459:395–433. [PMC free article] [PubMed]
28. Quadri L, Weinreb P, Lei M, Nakano M, Zuber P, Walsh C. Characterization of Sfp, a Bacillus subtilis phosphopantetheinyl transferase for peptidyl carrier protein domains in peptide synthetases. Biochemistry. 1998;37:1585–1595. [PubMed]
29. Christensen QH, Cronan JE. The Thermoplasma acidophilum LplA-LplB complex defines a new class of bipartite lipoate-protein ligases. J. Biol. Chem. 2009;284:21317–21326. [PMC free article] [PubMed]
30. Jiang Y, Chan CH, Cronan JE. The soluble acyl-acyl carrier protein synthetase of Vibrio harveyi B392 is a member of the medium chain acyl-CoA synthetase family. Biochemistry. 2006;45:10008–10019. [PubMed]
31. Gasteiger E, Gattiker A, Hoogland C, Ivanyi I, Appel RD, Bairoch A. ExPASy: the proteomics server for in-depth protein knowledge and analysis. Nucl. Acids Res. 2003;31:3784–3788. [PMC free article] [PubMed]
32. Hallmann C, van Aarssen BGK, Grice K. Relative efficiency of free fatty acid butyl esterification: Choice of catalyst and derivatisation procedure. J. Chromatog. A. 2008;1198-1199:14–20. [PubMed]
33. Laskey RA, Mills AD. Quantitative film detection of 3H and 14C in polyacrylamide gels by fluorography. Eur J Biochem. 1975;56:335–341. [PubMed]
34. Clarke L, Carbon J. A colony bank containing synthetic CoI EI hybrid plasmids representative of the entire E. coli genome. Cell. 1976;9:91–99. [PubMed]
35. Robinson NE, Robinson AB. Prediction of protein deamidation rates from primary and three-dimensional structure. Proc Natl Acad Sci U S A. 2001;98:4367–4372. [PMC free article] [PubMed]
36. Jordan SW, Cronan JE., Jr. Biosynthesis of lipoic acid and posttranslational modification with lipoic acid in Escherichia coli. Methods Enzymol. 1997;279:176–183. [PubMed]
37. Frottin F, Martinez A, Peynot P, Mitra S, Holz RC, Giglione C, Meinnel T. The proteomics of N-terminal methionine cleavage. Mol Cell Proteomics. 2006;5:2336–2349. [PubMed]
38. Xiao Q, Zhang F, Nacev BA, Liu JO, Pei D. Protein N-terminal processing: substrate specificity of Escherichia coli and human methionine aminopeptidases. Biochemistry. 2010;49:5588–5599. [PMC free article] [PubMed]
39. McMurry LM, Oethinger M, Levy SB. Triclosan targets lipid synthesis. 1998;394:531–532. [PubMed]
40. Hillis DM, Bull JJ. An empirical test of bootstrapping as a method for assessing confidence in phylogenetic analysis. Syst Biol. 1993;42:182–192.
41. Ferrari E, Henner DJ, Hoch JA. Isolation of Bacillus subtilis genes from a charon 4A library. J. Bacteriol. 1981;146:430–432. [PMC free article] [PubMed]
42. Wild J, Hradecna Z, Szybalski W. Conditionally amplifiable BACs: Switching from single-copy to high-copy vectors and genomic clones. Genome Research. 2002;12:1434–1444. [PMC free article] [PubMed]
43. Schulz GE. Binding of nucleotides by proteins. Curr. Opinion Struct. Biol. 1992;2:61–67.
44. Nesbitt NM, Baleanu-Gogonea C, Cicchillo RM, Goodson K, Iwig DF, Broadwater JA, Haas JA, Fox BG, Booker SJ. Expression, purification, and physical characterization of Escherichia coli lipoyl(octanoyl)transferase. Protein Expr Purif. 2005;39:269–282. [PubMed]
45. Posner MG, Upadhyay A, Bagby S, Hough DW, Danson MJ. A unique lipoylation system in the Archaea. Lipoylation in Thermoplasma acidophilum requires two proteins. FEBS J. 2009;276:4012–4022. [PubMed]
46. Doolittle RF. Convergent evolution: the need to be explicit. Trends Biochemi. Sci. 1994;19:15–18. [PubMed]
47. Omelchenko M, Galperin M, Wolf Y, Koonin E. Non-homologous isofunctional enzymes: A systematic analysis of alternative solutions in enzyme evolution. Biol. Direct. 2010;5:31. [PMC free article] [PubMed]
48. Wycuff DR, Matthews KS. Generation of an AraC-araBAD promoter-regulated T7 expression system. Anal.Biochem. 2000;277:67–73. [PubMed]
49. Cronan J. Cosmid-based system for transient expression and absolute off-to-on transcriptional control of Escherichia coli genes. J.Bacteriol. 2003;185:6522. [PMC free article] [PubMed]
50. Kouzminova EA, Kuzminov A. Chromosomal fragmentation in dUTPase-deficient mutants of Escherichia coli and its recombinational repair. Mol. Microbiol. 2004;51:1279–1295. [PubMed]
51. De Lay NR, Cronan JE. In vivo functional analyses of the type II acyl carrier proteins of fatty acid biosynthesis. J. Biol. Chem. 2007;282:20319–20328. [PubMed]
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